The enormous increase in the demand for synthetic oligonucleotides fueled by the advances in DNA technology over the past few decades has been accelerated by recent progress in sequencing and decoding whole genomes, particularly the human genome. A number of methods in molecular biology and DNA-based diagnostics to amplify, detect, analyze and quantify nucleic acids are dependent on chemically synthesized oligonucleotides. They are employed, for instance, in recombinant host-vector systems used for techniques such as site-directed mutagenesis. They are also employed in PCR methods, which use oligonucleotides as primers in temperature-cycled enzymatic amplifications of nucleic acids. Primers are also needed for state of the art sequencing techniques featuring enzymatic elongation and random termination. A rapidly growing field is the application of oligonucleotides in hybridization assays, which are based on the specific annealing of oligonucleotide probes to the region of a nucleic acid analyte having a complementary sequence. Probes with covalently conjugated dyes generating a fluorescent signal on perfect match hybridization are among the latest developments in this field. Corresponding methods to test for specific genomic epitopes, such as allelic discrimination or SNP detection, employ hybridization probes and are readily multiplexable. Thus, automated high-throughput systems capable of simultaneously screening a vast number of analytes using miniaturized arrays of hybridization probes, such as DNA chips, are used for applications such as genotyping and expression profiling. These and related methods may have an enormous impact in the areas of drug development and health management, as well as other fields.
Antisense technology is another field requiring a rapidly increasing supply of oligonucleotides. Synthetic antisense oligonucleotides are complementary to an RNA or DNA target sequence and are designed to halt a biological event, such as transcription, translation or splicing. They represent a whole new class of therapeutic agents, which have been shown to exhibit antiviral activity by inhibiting viral DNA or protein synthesis and moreover, may be able to cure certain diseases by inhibiting gene expression via specifically recognizing and binding mRNA. The most recent generation of antisense oligonucleotides comprise backbone-modified DNA or RNA derivatives such as 2′-OMe-RNA, phoshorothioate, morpholino nucleic acid, LNA, or combinations thereof. With these derivatives, the properties of antisense compounds, such as nuclease resistance, RNase H susceptibility and binding affinity, which are crucial for either blocking or promoting the degradation of the complementary mRNA, can be modulated and optimized.
Oligonucleotide primers with purities as low as 50% are usually sufficient for standard PCR applications and primers with purities of 70% can be effectively employed in sequencing methods. Other applications, however, generally require purities of greater than 90%. For example, contaminants in an oligonucleotide used as an insert in cloning experiments, after amplification in a host, may dominate the desired product nucleic acid due to their potentially better insertion properties. Antisense oligonucleotides intended for therapeutic purposes have to meet the high standards required by the FDA for drugs applied to humans. To reach purities greater that 90%, the crude product obtained after the cleavage/deprotection step at the end of the synthesis usually has to be subjected to subsequent purification steps, such as HPLC and/or PAGE, depending on the size and the sequence of a desired oligonucleotide.
The current state of the art in oligonucleotide synthesis is automated solid phase synthesis using phosphoramidite chemistry, which in particular is based on the developments of McBride et al. (1983) Tetrahedron Letters 24:245–248 and Sinha et al. (1983) Tetrahedron Letters 24:5843–5846, and which has been, together with related methods such as the hydrogen-phosphonate chemistry, extensively reviewed by Beaucage et al. (1992) Tetrahedron 48:2223–2311, each of which is specifically incorporated herein by reference in its entirety. During solid phase oligonucleotide synthesis, a series of nucleotide monomers are sequentially attached in a predetermined order to either, depending on the direction of chain extension, the 5′-functional group or the 3′-functional group of the growing oligonucleotide strand, which is linked to a solid phase such as CPG or a polystyrene resin. The method for attachment of each monomer is generally comprised of the following steps: 1) deprotection of the reactive functionality, usually the 5′-hydroxyl group, of the growing strand; 2) coupling by addition of a nucleoside monomer and an activator; 3) capping of unreacted terminal functional groups through introduction of an inert protective group, to prevent further coupling to failure sequences; and 4) oxidation of the newly formed internucleosidic phosphorous linkage to the naturally occurring pentavalent state.
This synthetic method has reached a high level of optimization featuring coupling efficiencies of up to 99% on average per cycle. Nevertheless, the small amount of non-extended oligonucleotide strands or failure sequences per cycle results in typical crude yields of about 50–80% for a 20-mer, with further reduced purities for longer oligonucleotides. Even with a highly optimized coupling efficiency of 99%, a 100-mer cannot be obtained in purities exceeding 35%. In the case in which failure sequences subsequently also fail to be capped during the capping step, they continue to participate in the following synthetic cycles. The resulting deletion sequences, ranging from (n−1)-mers and (n−2)-mers to shorter lengths, are present as impurities in the crude full-length oligonucleotide (n-mer). Among these, (n−2)-mers and shorter contaminants are readily removable by chromatographic purification steps, such as HPLC using reverse or anion-exchange phases. However, these methods are unsuitable for separating the (n−1)-mer impurities from the desired (n)-mer. Purification can be achieved by applying PAGE electrophoresis, if the respective low recovery rates are acceptable.
Thus, an effective capping process during the course of every synthetic cycle of an oligonucleotide synthesis is essential for inhibiting failure sequences from undergoing further elongation. In other words, capping allows for the truncation of failure sequences thereby largely reducing the formation of deletion sequences. Since the presence of deletion sequences, having almost identical size and composition as the product, makes purification very difficult, the main utility of capping is to minimize the length and presence of failure sequences.
In current state of the art methods, capping during the course of solid phase phosphoramidite oligonucleotide synthesis is accomplished by acetylation of the 5′-ends of failure sequences employing acetic anhydride in the presence of N-methylimidazole, as described by Eadie et al. (1987) Nucleic Acids Res. 15:8333–8249, which is specifically incorporated herein by reference in its entirety. For oligonucleotide synthesis using the hydrogen-phosphonate approach, Andrus et al. (1988) Tetrahedron Letters 29:861–864, have introduced phosphite monoesters as capping reagents, thus exploiting the coupling chemistry also for the capping reaction. Accordingly, Yu et al. (1994) Tetrahedron Letters 35:8565–8568 (incorporated herein by reference in its entirety), have described the use of diethoxy N,N-diisopropyl phosphoramidite as a capping reagent for a phosphoramidite-based oligonucleotide synthesis. Both of the latter capping methods provide for caps that survive the final cleavage and deprotection of the synthesized oligonucleotide, which is advantageous for applications involving enzymatic reactions capable of excluding the undesired 5′-capped failure sequences e.g. from getting inserted into a vector.
Various strategies comprising capping reagents providing for post-synthetic removal of truncated sequences have been described. For example, the use of lipophilic capping reagents, enabling the separation of capped sequences from the full-length sequences on a hydrophobic stationary phase has been described by Natt et al. (U.S. Pat. No. 6,107,479 and Tetrahedron 53:9629–9636 (1997)). Since this purification step is based on unspecific, non-covalent interactions it relies on time-consuming chromatographic techniques, which are hardly suited for automation. Thus, it is not a substantial improvement over standard procedures, for instance RP-HPLC purification subsequent to processing the oligonucleotide synthesis without removal of the DMT-group in the final synthetic cycle (‘trityl-on’). Coolidge et al., U.S. Pat. No. 5,464,759, have described the use of capping reagents bearing an antigenic site. Following oligonucleotide synthesis and cleavage from the solid support, the crude product is eluted through a resin that is derivatized with the corresponding antibodies. Due to the specific immunological interaction, the truncated contaminants are removed and a purified product is obtained. Coolidge et al. do not convincingly teach, however, how to bind the cap including the antigenic site to the failure sequences in such a manner that it can survive the final cleavage/deprotection step. Both deprotection and intact caps are crucial for the post-synthetic purification step intended by the authors. Furthermore, the application of antibodies requires highly skilled personnel and lacks cost efficiency.
Coolidge et al., have also suggested the post-synthetic immobilization of contaminants using capping reagents comprising a functionality that is able to covalently bind to a correspondingly derivatized matrix. Coolidge et al., however, do not teach how to accomplish this method of purification.